Through the development of recombinant DNA techniques, it has become fairly straightforward to clone DNA sequences from essentially any organism into plasmid or viral vectors for propagation and amplification in a foreign host. In this form the DNA can be studied with regard to its sequence, structure, coding capacity or other properties. It can also be used for a variety of applications such as detection of complementary sequences in samples and the generation of altered forms of a gene product.
One method for producing altered forms of a gene is known as site-specific mutagenesis. Site-specific mutagenesis is a term used to denote the generation of specific base substitutions at selected sites in the DNA. Site-specific mutagenesis is a valuable tool for the study of DNA function and protein structure and function. A number of different mutagenesis methods have been reported (Smith, M., 1985, Ann. Rev. Genet. 19, 4233; and Section IV, Chapters 17-21, 1987, Meth. Enzymol., 154, 329-414). Hutchison. et al. (1978, J. Biol. Chem., 253, 6551-6560) introduced a method to obtain site-specific changes in DNA sequences using single-stranded DNA (ssDNA) and a synthetic oligonucleotide. The oligonucleotide is complementary to the single-stranded template DNA except for a region of mismatch in the center. It is this region that contains the desired nucleotide change or changes. According to Hutchison, et al., the general method for obtaining site-specific changes in a DNA sequence is as follows. The synthetic oligonucleotide is hybridized to the ssDNA. This mismatched hybrid (heteroduplex) serves as a template for the enzymatic synthesis of a complementary mismatch (mutant) strand. Following hybridization, the oligonucleotide is extended with DNA polymerase to create a double-stranded structure. The nick is then sealed, and the resulting heteroduplex is transformed into Escherichia coli (E. coli) host. Upon DNA replication and strand segregation, the cell contains a mixture of wild-type and mutant templates. Because mutant and wild-type plasmids are present in the same cell, a second round of transformation is generally employed to insure genetic purity. This method of in vitro mutagenesis is generally employed using single-stranded M13 or phagemid templates. Although the yield of mutants should theoretically be 50%, in practice the yield is much lower. Contributing factors, such as incomplete in vitro polymerization, primer displacement by a DNA polymerase used in the fill-in reaction and in vivo host-directed mismatch repair mechanisms, which favor repair of unmethylated newly synthesized DNA strands (Kramer, B. et al., 1984, Cell, 38, 879), contribute to the lower yield.
In order to increase the frequency with which the desired mutation is isolated, a number of selection techniques have been described. These selection techniques are generally directed to a method for constructing a mutation in DNA by hybridizing a section of parent DNA with a synthetic oligonucleotide which is mostly complementary to the parent DNA strand, but has one or more base pair mismatches at the desired point of mutation. This hybridized DNA strand is transformed into a bacterial host where the hybridized DNA strand can replicate, with the strand having the mismatch serving as the template for the desired mutation. The problem that exists at this point is that mutant and non-mutant strands of DNA are present and the mutant strand must be isolated.
Kramer, et al. (1984, Nucleic Acids Res. 12, 9441-9456) describe a method for introducing mutations into recombinant genomes of filamentous phage M13 ssDNA. The method involves the construction of a double-stranded "gapped duplex" DNA where the (+) longer strand parent DNA has two amber mutations which prevent it from replicating in a non-suppressing host. An "amber mutation" is a class of suppressible mutations that results in the creation of a UAG codon in mRNA. This codon normally signifies translation termination, so that polypeptide synthesis stops at the amber site. The shorter (-) DNA strand has the two amber mutations removed. When a mismatched synthetic oligonucleotide is hybridized to the single stranded DNA in the gap of the gapped duplex and connected with the rest of the (-) strand, the newly constructed (-) strand can replicate in a non-suppressing host. For this reason, the parent (+) strand with the amber mutations can be selected against the newly constructed (-) strand. Kramer, et al. report mutant recovery of 70 percent or more. A drawback of this system is that the complementary strand of the restriction fragment can compete with the ssDNA for hybridization, and the reannealed fragment can create a background of non-mutant plaques.
Another selection strategy is based on the method of Kunkel (1985, Proc. Natl. Acad. Sci. 82, 488-492) and Kunkel, et al. (1987 Methods Enzymol. 154, 367-382). In this scheme ssDNA is prepared in a special E. coli host which is dut.sup.- ung.sup.-. This phenotype results in the occasional substitution of uracil for thymidine in the DNA strand. Mutagenesis is performed in the usual manner by hybridizing a mismatched oligonucleotide to the ssDNA template and filling in the outside strand with DNA polymerase. When this molecule is transformed into a dut.sup.+ ung.sup.+ E. coli strain, the inside strand containing the uracil substitutions is cleaved and destroyed, leaving only the mutant strand intact. However, a low number of transformants is generally obtained.
Variations of the in vitro mutagenesis technique using two oligonucleotide primers have also been described (Zoller, M. J. and Smith, M. Methods in Enzymology (1987) Vol. 154, 329-351) and have been used without either of the oligonucleotides conferring a selectable phenotype (Norris, et al. 1983, Nucleic Acids Res. 11, 5103-5113).
Carter, et al. (1985, Nucleic Acids Res. 13, 4431-4443) describe a method for the construction of mutations in M-13 vectors using synthetic oligonucleotides whereby the DNA is first cloned into an engineered M13 vector which carries a genetic marker so that it can be selected against the parent strand. The technique used is referred to as "coupled priming" where one oligonucleotide with base-pair mismatches is hybridized to the parent DNA strand to construct the mutation of interest, and a second oligonucleotide which contains a selectable marker, is also hybridized to the parent strand. The two oligonucleotides are connected to make a continuous strand of DNA. This heteroduplex DNA is transfected into a mismatch repair deficient strain of E. coli which can select against the parent strand. In this case the primary marker used is an EcoK or EcoB marker which will cause the strand to be cleaved if transfected into an organism with that restriction enzyme. For example, by transfecting into a host organism that has the EcoK restriction enzyme, one can select against a parent strand containing an EcoK site, which will be cleaved. Because the EcoK marker can be changed to an EcoB marker by changing just one base pair, it is possible to cycle between the two markers for successive rounds of mutation by simply hybridizing a marker strand with the one base pair change to the parent strand. Mutant yields of up to 70 percent were reported with this process. However, it is unlikely that this method will work if the section of DNA cloned into the M13 vector contains an EcoK or EcoB site, since that site would also be cleaved.
Stanssens et al. (1989, Nucleic Acids Res. 17, 4441-4454) are directed to a method of construction of multiple mutations in a sequential manner through oligonucleotide-directed mutagenesis. Unlike Kramer, et al. and Carter, et al., Stanssens, et al. construct mutations in plasmids, which are doubled-stranded rather than in M-13 phage vectors, which are single-stranded. The Stanssens, et al. method utilizes the Kramer, et al. method of mutation insertion into a gapped-duplex DNA. In order to construct the gapped-duplex DNA, it is necessary to take double-strand plasmid DNA and turn it into ssDNA. This is accomplished because the plasmid used has had the origin of replication for the filamentous phage f1 (f1 replication origin) engineered into it, and this can be used to generate ssDNA. The orientation of the f1 DNA determines which of the two strands of the plasmid will be secreted. This allows one directional copying of one strand of the plasmid DNA. The chimeric ssDNA phage-plasmid vector containing a phage replication origin is known as a "phagemid."
In the Stanssens, et al. method, the gapped-duplex DNA is constructed using two complementary plasmids. One contains an ampicillin resistance gene and a chloramphenicol resistance gene that has been inactivated with an amber mutation, making the plasmid chloramphenicol sensitive; and a second plasmid that is chloramphenicol resistant and has an ampicillin resistance gene inactivated with an amber mutation. Antibiotic resistance is used to select for the mutant strands of DNA. Because the two plasmids have opposite antibiotic resistances, by alternating construction of the gapped duplex, additional mutations can be inserted in successive rounds of mutation and one may alternately select for the mutant strand. Both plasmid vectors contain a multilinker region which provides an area for insertion of target sequences for mutagenesis. One disadvantage of this technique is that hybridization of the antibiotic resistance restoration strand is inefficient because of its source as a double-stranded restriction fragment, the other strand which competes with the ssDNA for hybridization. In addition, reformed double-stranded restriction fragments will create a background of ampicillin resistant non-mutant colonies, limiting the efficiency of the method.
Two other methods have been described for the selection of the mutant strand following the fill-in reaction in oligonucleotide directed in vitro mutagenesis. Vandevar, et al. (1988, Gene 65, 129-133) performed the fill-in reaction using 5-methyl dCTP to form a hemimethylated duplex strand. Nicks are introduced selectively into the nonmethylated parent strand through the action of the restriction endonuclease Msp I. The parent strand is then degraded through the action of exonuclease III. In a similar system sold by Amersham Corp., the in vitro fill-in reaction following hybridization of a mismatched oligonucleotide to a ssDNA template is performed using alpha-thio dCTP. The non-phosphorothioate non-mutant strand is then nicked specifically with the restriction enzyme Nci I and then degraded by the action of exonuclease III. Mutagenesis efficiencies of up to 95% are reported in this system. However, a drawback is the relatively large amount of DNA needed (up to 20 micrograms (ug)) to perform a reaction and transformation. Further, the system requires a series of complex enzymatic reactions.